This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Formula display:

Abstract

The repressor protein PhaR, which is a component of poly[(R)-3-hydroxybutyrate] granules, functions as a repressor of the gene expression of
the phasin PhaP and of PhaR itself. We used a quartz crystal microbalance to investigate
the binding behavior by which PhaR in Ralstonia eutropha H16 targets DNAs and amorphous poly[(R)-3-hydroxybutyrate] thin films. Binding rate constants, dissociation rate constants,
and dissociation constants of the binding of PhaR to DNA and to amorphous poly[(R)-3-hydroxybutyrate] suggested that PhaR bind to both in a similar manner. On the
basis of the binding rate constant values, we proposed that the phaP gene would be derepressed in harmony with the ratio of the concentration of the target
DNA to the concentration of amorphous poly[(R)-3-hydroxybutyrate] at the start of poly[(R)-3-hydroxybutyrate] synthesis in R. eutropha H16.

Keywords:

Introduction

Polyhydroxyalkanoate (PHA), an eco-friendly and biodegradable polyester, is synthesized
by a variety of bacteria, as their intracellular storage material for carbon and energy
(Doi et al. 1995; Steinbuchel and Fuchtenbusch 1998; Sudesh et al. 2000). In bacterial cells, PHA forms granules that are covered with a layer composed of
proteins and phospholipids (Potter et al. 2002). The most abundant constituent of this layer is phasin (PhaP). The presence of PhaP
on the surface of PHA granules contributes to the reduction in size of PHA granules
as well as to the slight enhancement of PHA production (Kojima et al. 2006; Potter et al. 2002; Potter and Steinbuchel 2005). Recently, the ability of PhaP to bind to a hydrophobic surface was used to develop
methods for protein purification, drug delivery, and tissue engineering applications
in in vitro experiments (Backstrom et al. 2007; Banki et al. 2005; Wang et al. 2008). In the cells of microorganisms, a repressor protein PhaR regulates the expression
of phaP and phaR. PhaR has also been reported to sense the presence of PHA and to interact with nascent
PHA granules, resulting in the derepression of phaP expression (Potter et al. 2002; Potter and Steinbuchel 2005). The presence of genes homologous to PhaR and PhaP in the genomes of various PHA-producing
bacteria suggests that a similar regulatory system by PhaR is likely to exist in PHA-producing
bacteria (Eugenio et al. 2010; Kojima et al. 2006; Maehara et al. 2002; Yamada et al. 2007; Yamashita et al. 2006). This regulatory system of PHA production through phaR and phaP expression can be applied in a two-hybrid system for protein-protein interaction
(Wang et al. 2011). Therefore, understanding of the regulatory system provides meaningful benefit to
not only basic science but also applications in various fields such as industry and
medicine.

In previous studies, the binding behaviors of PhaR to target DNA (including the promoter
region of phaP) and to melt-crystallized thin films of poly[(R)-3-hydroxybutyrate] [cr-P(3HB)] were investigated using surface plasmon resonance
(SPR) and quartz crystal microbalance (QCM) measurements (Yamada et al. 2007; Yamashita et al. 2006). However, kinetic parameters such as the binding rate constant (kon) and dissociation rate constant (koff) by which PhaR targets DNA and P(3HB) have not been determined thus far. These kinetics
and stoichiometric analyses will contribute new insights into the behavior of PhaR
in the regulatory system of phaP expression. In order to determine the precise kinetic parameters, we selected a multichannel
QCM sensing system to monitor the binding reaction of PhaR from Ralstonia eutropha H16 to target DNAs (including the promoter regions of phaP and phaR) and thin films of amorphous P(3HB) [am-P(3HB)] derived from atactic P(3HB). This
is because the P(3HB) native granule is composed of am-P(3HB). Recently, the regulatory
system of PHA production through phaR and phaP expression has been applied in studies of protein-protein interaction, protein purification,
drug delivery, and tissue engineering. The insights gained into this regulation mechanism
in this study have the potential to improve applications in white biotechnology. We
have determined kinetic parameters based on mass changes on the DNA-immobilized and
am-P(3HB)-coated QCM oscillators, and discuss the binding behavior of PhaR with target
DNA and am-P(3HB).

Methods

Expression and purification of autoregulator protein PhaR

All chemical reagents were purchased from Wako Pure Chemicals (Osaka, Japan). The
phaR gene from R. eutropha H16 was cloned using a TOPO TA cloning Kit (Invitrogen, Carlsbad, CA) with the forward
primer 5′-CACCATGGCCACGACCAAAAAAGG-3′ and reverse primer 5′-TTACTTCTTGTCCGGCTGGT-3′.
The resultant plasmid is referred to as pET100/D-TOPO-PhaRRe. The expression of the phaR gene was driven by the T5 promoter, which is inducible with isopropyl-α-D-thiogalactopyranoside (IPTG). The constructed plasmid was introduced into Escherichia coli BL21(DE3). Transformants were grown in 1200 mL of Luria-Bertani medium containing
ampicillin (100 μg/mL) and kanamycin (50 μg/mL). They were cultivated at 30°C until
the OD600 of the culture reached 0.5. After the addition of IPTG (final concentration of 1
mM), the transformants were grown for an additional 5 h. The cells were then harvested
and washed with chilled buffer A (50 mM sodium phosphate (pH 8.0) containing 300 mM
NaCl and 10 mM imidazole), and were suspended in 60 mL of the same buffer. The suspension
was stored at −80°C until use. The suspension was thawed on ice and disrupted by sonic
oscillation, also on ice. The cell debris was then removed by centrifugation at 15000 × g for 60 min at 4°C, and the supernatant was collected for purification. The experiments
were carried out at 4°C throughout the purification steps. The crude extract was shaken
gently with nickel-nitrilotriacetic acid agarose (Qiagen, Valencia, CA) for 1 h. The
mixture was then poured into a column. The column was washed with buffer A containing
20 mM imidazole, and then the His-tagged protein was eluted with buffer A containing
250 mM imidazole. The eluates containing PhaR were dialyzed against 10 mM HEPES (pH
7.4) containing 150 mM NaCl and 3 mM EDTA and stored at −80°C. The protein concentration
was determined using a Bio-Rad Protein Assay Kit (Bio-Rad, Hercules, CA) with bovine
serum albumin as the standard. Proteins were separated by sodium dodecyl sulfate (SDS)-12.5%
polyacrylamide gel electrophoresis (PAGE) and stained with Coomassie brilliant blue
(CBB) R-250 (BioRad) as described by Laemmli (Figure 1).

Calibration of 27-MHz QCM in aqueous solution

The QCM apparatus was an AFFINIX Q4 (Initium Co., Ltd., Tokyo, Japan) with 4 500-μL
cells equipped with a 27-MHz QCM plate (8.7 mm diameter quartz plate and 5.7 mm2 area Au electrode) at the bottom of the cell and a stirring bar with a temperature
control system (Takahashi et al. 2007; Takahashi et al. 2008). The relationship between mass and frequency changes in aqueous solutions when DNAs
and/or proteins were immobilized onto the QCM was calibrated by comparing it against
values in the air phase. One Hz of frequency represents a 0.10 ng cm-2 mass increase on the QCM plate. The noise level of the 27-MHz QCM was ±2 Hz in buffer
solutions at 25°C, and the stability of the frequency was ±2 Hz for 1 h in buffer
at 25°C.

Preparation of the DNA-Immobilized QCM Oscillator

The structures of the biotinylated oligonucleotides used in this study are summarized
in Table 1: they consisted of 5-biotinylated dsDNA (50 bp) containing a site recognized by PhaR
dsDNA (phaP promoter region DNA and phaR promoter region DNA) and no-site dsDNA (control DNA). Oligonucleotide duplexes were
formed by mixing a biotinylated strand and its complementary strand in a solution
of 10 mM Tris–HCl (pH 7.8), 1 mM EDTA, and 200 mM NaCl, and then boiling for a few
minutes, followed by cooling to room temperature over 1 h. These oligonucleotides
were immobilized on a cleaned Au electrode of the QCM using biotin-avidin linkage
according to the methods described in a previous paper (Okahata et al. 1998). The amount of immobilized DNA was maintained at 191 ng (0.55 – 0.02 pmol) cm-2, which corresponds to 1% coverage of the Au surface (5.7 mm2). This would allow sufficient space to accommodate the binding of a large enzyme
molecule.

Table 1.Kinetic parameters for the binding of PhaR to DNAs and P(3HB) on the 27-MHz QCMa

Preparation of P(3HB) thin films

QCM oscillators were washed with a freshly prepared Piranha solution of H2O2/H2SO4 (1/3 v/v) and were rinsed several times with Milli-Q water. (Caution: Piranha solution
is very oxidative and dangerous, and direct contact should be avoided). Thin films
of P(3HB) were prepared on the QCM oscillators by casting 300 μl of chloroform solutions
(1.0–1.5 wt%) of the polymers on a spin-coater at 4000 rpm under dry air.

Reactions in the DNA-immobilized or am-P(3HB) coated QCM oscillator

Enzyme reactions in a DNA-immobilized or am-P(3HB) coated QCM cell were performed
with 500 μL of assay buffer (10 mM HEPES (pH 7.4), 150 mM NaCl, and 0.002% Tween 20).
The frequency changes in response to the addition of enzymes were then followed over
time. The solution was vigorously stirred to avoid any effects from the slow diffusion
of the enzymes. The stirring did not affect the stability or magnitude of the frequency
changes.

Results

In order to measure the binding behavior of PhaR to 5′-biotinylated dsDNAs (50 bp),
the DNA fragments with phaR-binding sequences (the promoter regions of phaR and phaP) and a non-specific sequence (negative control) were immobilized on the electrode
of a QCM by biotin-avidin linkage, according to methods outlined in previous papers
(Matsuno et al. 2001; Okahata et al. 1998). PhaR was purified using the His-tag purification system, and the purity of PhaR
was confirmed by SDS-PAGE (Figure 1). The binding behaviors of PhaR to the DNA fragments were monitored. Figure 2A shows a typical frequency decrease (mass increase) as a function of time, in response
to the addition of PhaR. PhaR mainly bound to the DNA containing the phaP promoter region (curve a), and barely bound to the DNA containing the phaR promoter region (curve b) and the control DNA (curve c). Figure 2B shows that the amount of the bound PhaR (Δm) followed a saturation curve as a function of the PhaR concentration ([PhaR]). These
binding curves formed a sigmoid curve.

Next, we attempted to determine the kinetic parameters (kon, koff, and Kd values) of the binding of PhaR to DNAs. The process by which PhaR binds to the DNAs
is described by Equation 1 (Okahata et al. 1998). The amount of the DNA/PhaR complex ([DNA/PhaR]) formed after the injection is given
by Equations 2 and 3. The fitting curves of the decreases in frequency at various
PhaR concentrations gave the relaxation time (τ) and the relaxation rate (τ-1) of PhaR binding. When the concentration of PhaR increased from 2.5 to 10 nM, the
amount of PhaR bound to the DNA increased (Figure 3A). In addition, the τ value decreased with the concentration of PhaR. The 1/τ value
at each PhaR concentration was plotted against the concentrations of PhaR, according
to Equation 4 (Figure 3B). The PhaR binding and dissociation rate constants (kon and koff) were obtained from the slope and the intercept of Equation 4, respectively. The
Kd values were obtained from the ratio koff/kon. The kinetic parameters, kon, koff, Kd, for the target DNAs and control DNA are summarized in Table 1.

(1)

(2)

(3)

(4)

Figure 3.The influence of PhaR concentrations on the rate of binding to target DNAs. (A) Binding behaviors of PhaR to the DNA including the phaP promoter region in response to changes in PhaR concentration. The arrow indicates
the time of enzyme injection. (B) Linear reciprocal plots of the relaxation rate (τ-1) against the PhaR concentrations according to Eq. (4) in the text.

We also investigated the binding of PhaR to P(3HB) granules in microorganisms using
a QCM sensor coated with am-P(3HB) (Figure 4A). This is because the P(3HB) native granules are mainly composed of am-P(3HB). The
amount of PhaR bound on am-P(3HB) thin films increased when the concentration of PhaR
increased from 1 to 15 nM (Figure 4B and C). Interestingly, as with the binding curve against the target DNA, the binding
curve against am-P(3HB) exhibited a sigmoid curve. These results indicated that PhaR
bound to P(3HB) in a similar manner as to DNA. The kinetic parameters (kon and koff) were calculated from Equations 1 to 4 (Figure 4D and Table 1). The kon value for am-P(3HB) (kon = 7.0 ± 3.8 × 104 M-1 s-1) showed no significant difference from that for DNA containing the phaP promoter region (kon = 6.0 ± 0.4 × 104 M-1 s-1).

Figure 4.Binding behavior of PhaR to am-P(3HB) thin films. (A) Typical time courses of the frequency changes of am-P(3HB) coated on a QCM oscillator,
in response to the addition of PhaR. (B) Saturation binding of PhaR to P(3HB). The curve was fitted with a sigmoid curve.
[P(3HB)] = 200 μg cm-2 on a QCM, in 10 mM HEPES buffer solution (pH 7.4), 150 mM NaCl, and 0.001% Tween
20 at 25°C. (C) Binding behavior of PhaR to P(3HB) in response to change in the PhaR concentration.
The arrow indicates the time of enzyme injection. (D) Linear reciprocal plots of the relaxation rate (τ-1) against PhaR concentrations according to Eq. (4) in the text.

Discussion

In order to understand the regulatory system governing PHA production in detail, we
investigated the binding behaviors of PhaR to the target DNA (containing the promoter
region of phaP) and to P(3HB), using QCM measurements. Regarding PhaR-DNA binding, Figure 2A shows that PhaR mainly bound to the DNA containing the phaP promoter region (curve a), and barely bound to the DNA containing the phaR promoter region (curve b) or to the control DNA (curve c). The binding curve of PhaR
to the phaP promoter region showed sigmoid curve, implying that PhaR binds to target DNA in a
cooperative reaction. The SPR analysis of PhaR-DNA binding in previous studies was
not capable of monitoring the initial binding of PhaR, because the concentration of
PhaR (10 μM) was higher than in the present experimental conditions (2.5 to 10 nM)
(Kojima et al. 2006; Maehara et al. 2002). The higher binding affinity of PhaR to the phaP promoter region accorded with the results of gel-mobility-shift assays (Maehara et
al. 2002). The DNA fragments with the phaP promoter region shifted at a lower concentration of PhaR compared to the DNA fragments
that contained the phaR promoter region (Potter et al. 2002).

The binding rate constant for the DNA containing the phaR promoter region (kon = 0.5 × 104 M-1 s-1) was similar to the parameters for the control DNA (kon = 0.4 × 104 M-1 s-1) (Table 1). Moreover, the dissociation rate constant for the DNA containing the phaP promoter region (koff = 1.7 ± 0.4 × 10-3 s-1) was not significantly different from the dissociation rate constant for the DNA
containing the phaR promoter region (koff = 0.9 × 10-3 s-1) or that of the control DNA (koff = 0.7 × 10-3 s-1). These parameters indicate that PhaR had higher affinity for the phaP promoter region than for the phaR promoter region. The larger kon value for the DNA with the phaP promoter region must have been due to the length of the recognition sequence for
PhaR in the target DNA region. A 32-bp region TGC-rich sequence is recognized by PhaR
in the phaP promoter region, while the phaR promoter region included only an 8-bp recognition sequence (Table 1) (Potter and Steinbuchel 2005). On the basis of the kon values obtained in this study, the difference in kon values between the promoter regions of phaP and phaR corresponds to the hypothetical model of PhaR-mediated phaP expression (Maehara et al. 2002; Potter and Steinbuchel 2005; Yamada et al. 2007). In particular, when PHA is not accumulated in the cells, the presence of PhaR is
necessary to repress the gene expression of phaP. Since PhaP is a predominantly PHA granule-associated protein, PhaP production is
not required for the cells without PHA accumulation (Maehara et al. 2002). Thus, the lower kon value for DNA with the phaR promoter region indicates weak repression of phaR expression in cells.

In the measurement of PhaR-am-P(3HB) binding, we did not obtain koff and Kd values of the binding of PhaR to am-P(3HB), because the koff was negative. This result indicates that the binding of PhaR to am-P(3HB) is an irreversible
interaction (Table 1). There was no significant difference between the kon value for am-P(3HB) (kon = 7.0 ± 3.8 × 104 M-1 s-1) and that for the DNA containing the phaP promoter region (kon = 6.0 ± 0.4 × 104 M-1 s-1), which implied that the derepression of phaP expression was prompted by an increase in the concentration of am-P(3HB) in the cells.
In other words, the concentration-dependent effect was one of the main factors initiating
the expression of phaP at the onset of the dissociation of PhaR from the phaP promoter region in cells.

In conclusion, we observed initial binding behaviors between PhaR and target molecules
such as target DNAs and am-P(3HB), using QCM techniques. Based on the QCM data, kinetic
parameters (kon, koff, and Kd) for the binding of PhaR to target molecules were determined by the kinetic analysis
of obtained binding curves. These values provided a novel insight into the binding
behavior of PhaR with target molecules. The phaP gene is likely derepressed in harmony with the ratio of the concentration of the
target DNA to the concentration of am-P(3HB) at the beginning of P(3HB) synthesis
in microbes. On the basis of the results of a previous paper (Maehara et al. 2002), we assumed that PhaR dissociates from the PhaR/DNA complex when P(3HB) is accumulated
under intracellular conditions. This finding indicates that the effector molecules
of PhaR are P(3HB) molecules. Also, one of the factors responsible for the dissociation
of PhaR from DNA is the high affinity of PhaR to P(3HB). The binding of PhaR to DNA
and to am-P(3HB) showed similar kon values, suggesting that a concentration-dependent effect caused the expression of
phaP with dissociation of PhaR from the phaP promoter region. The insights of the regulation mechanism concerning PhaR in PHA
synthesis have the potential to improve the applications of PHA in white and red biotechnology.